Flow cytometry is a technique for analyzing individual cells as they flow in a liquid through a beam of light. When cells passes through the light, several sensors detect
25 light scattering and fluorescence, and register properties of the individual cell. The light signals are then converted to electronic signals that can be analyzed on a computer. This allows measurement of both physical properties of cells, including size and granularity, as well as fluorescence signal within the cell. Several
wavelengths of fluorescence can be measured simultaneously and allow analysis of many components within a single sample. Cells can be analyzed by staining with fluorescent dyes that bind directly to the component(s) of interest, or through
antibodies conjugated to a fluorescent dye. Cells are often „fixated‟ prior to staining, as it prevents autolysis and degradation of cell components. Fixation also
permeabilizes cellular membranes and allows dyes or antibodies to penetrate the cell.
Flow cytometry allows high-rate analysis of individual cells, and is a useful tool for investigating cell properties that cannot be detected by other methods, including viability assays. Although each cell that passes through the sensor is analyzed, data is commonly presented in fraction of the total number of cells analyzed. Flow
cytometry was used during this master thesis to investigate the effects of treatment on cell cycle distribution and apoptosis. Analysis by flow cytometry was done in collaboration with Idun Dale Rein at the Flow Cytometry Core Facility (Cancer Research Institute, Radium Hospital, Norway).
Cell fixation protocol
Cells were treated as previously described and incubated in T25 nuncleon flasks.
After 24 hours of incubation, cells were detached by trypsin-EDTA, washed with PBS, and centrifuged at 1700 g for 3 minutes. PBS was removed, and cell pellets were permeabilized by drop-wise addition of ice-cold methanol (VWR Chemicals).
Cells were kept at - 20ºC for minimum 24 hours to ensure permeabilized cell membranes. In addition, one sample was extracted prior to treatment (at 0 hours), fixated as described above and stained together with samples from the same
experiment.
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3.6.1 TUNEL assay
Terminal deoxynucleotidyl transferase (TdT) dUTP Nick-End Labeling (TUNEL) assay is used to detect apoptotic cells by incorporation of labeled nucleotides by TdT enzymes. Cells that undergo apoptosis activate endonucleases that degrade and form „nicks‟ within DNA. These „nicks‟ within the DNA strand can be filled in with labeled nucleotides by TdT enzymes. The TUNEL assay uses dioxyuridine
Triphosphate (dUTP) molecules conjugated to biotin as substrate for the TdT enzyme. If the biotinylated UTP is incorporated into the DNA of a cell, it can be detected by addition of labeled-streptavidin, which has a high specific affinity for biotin. The labeled streptavidin used for TUNEL assay contained cyanine 5 (Cy5) – which emits fluorescent light at peak 650 nm.
Protocol
Prior to staining, fixated cells were transferred to 5 mL Falcon® round-bottom polystyrene test tubes (Corning Inc.) and washed with PBS. The TUNEL-assay reaction mixture was prepared according to manufacturer, and 40 µl of the mixture was added to each sample which were incubated at 37ºC for 30 minutes. 3 ml of PBS was then added to each sample, and the samples were centrifuged at 1000 g for 3 minutes and the PBS was removed by pipette. Dry milk was mixed with PBS to a 5% weight/volume ratio and centrifuged at 2000 g for 3 minutes. The supernatant was used to dilute the Cy5-labeled streptavidin to a 1:400 concentration. 100µL of the solution containing streptavidin (1:400) was then added to each sample, and the samples were covered in aluminum foil and incubated for 40 minutes. 3 mL PBS was added after
incubation, samples were centrifuged at 1700 g for 3 minutes, and PBS was removed. Finally, samples were stained with Hoechst before flow cytometry analysis.
3.6.2 Cell cycle analysis
Analyzing cell cycle distribution can be useful for investigating cellular changes that occur after a specific treatment. Cell cycle analysis is based on the principle that cells replicate their DNA as they move through the cycle. When DNA is replicated
27 the total amount of DNA within a cell increases from n to 2n, which can be
measured and used to analyze the distribution of cells within the G1-, S- and G2 /M-phase by flow cytometry. In this study, cells were stained with Hoechst 33258
(Invitrogen) to determine the cell cycle distribution of cells after treatment. Hoechst is a nucleic acid stain that binds double-stranded DNA and emits blue fluorescence that can be detected by flow cytometry.
Protocol
Prior to staining, fixated cells were transferred to 15 mL round-bottom centrifuge tubes and washed with PBS. For convenience, samples were always stained by TUNEL-assay before Hoechst. Hoechst 33258 was diluted in PBS (1:400) and 300-600 µL was added to each cell pellet, depending on the amount of cells present.
Samples were kept at 4ºC overnight before flow cytometry analysis. Before flow cytometry, cell samples were passed through the Falcon® cell strainer cap (Corning Inc.).
3.6.3 Flow cytometry: gating strategy and data analysis
Analysis of flow cytometry data was done with the FlowJo™ software. Regardless of staining method, gating is required to exclude any aggregated or fragmented cells to allow analysis of single, stained cells. For this study, the Hoechst DNA stain was used to gate for single cells within each sample by including only cells with a cell-volume (area) which corresponded to the single-celled population (figure 5A).
Further analysis was then made within the single cell population. When single cells had been gated, apoptosis and cell-cycle analysis could follow. TUNEL- analysis was made by creating gates around cells with little or no Cy5 stain (figure 5B).
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Figure 6: Gating strategy of B76 cells stained by Hoechst and TUNEL afte r flow cytometry. A) Gating of single cells within the total sample removes unwanted doublets, B) analyzing single cells by Cy5 staining from TUNEL assay is used to estimate the non -apoptotic/apoptotic fraction within the sample, C) Histogram of the single-celled population by DNA content, D) Cell cycle distribution as analyzed in FlowJo using the „Watson‟ algorithm.
Although this method is less accurate, consistent gating increases precision and gives good indication of apoptotic fractions within each sample. The fraction of apoptotic cells shown within the single-cell observations in figure 6A, for instance, was estimated to be 4.4 % (figure 6B). Cell cycle analysis of Hoechst stained cells was done using the integrated cell-cycle analysis tool in FlowJo. The cycle
distribution is calculated by a Watson „pragmatic‟ algorithm which estimates the fraction of cells in G1 and G2/M-phase. The model assumes a normal distribution of
29 cells within the G1 and G2 phase, and uses these values to estimate the frequency of cells in S-phase. Cell cycle analysis may require optimization if samples contain noise or have broad G1 and G2 peaks. The model „fit‟ is reported as root mean square (RMS), where RMS values <1.5 are generally considered as „good‟ fit.